Preparation of Material for TEM Examination



Brendan Choat, Steven Jansen, Chrissie Prychid


This protocol covers the procedure for fixation, embedding and staining of material for examination by transmission electron microscopy (TEM). This includes working with various embedding resins, formvar grids and immunogold labelling.


Transmission electron microscopy can produce very high resolution (sub nm) images from ultrathin sections of plant tissue. Material is typically fixed and then embedded in resin before being sectioned with an ultramicrotome. Embedding and sectioning procedures are typically laborious and time-consuming, often involving dangerous compounds. However, because of the high resolution, TEM can often reveal structural information not available with light microscopy or scanning electron microscopy.


  • Fume hood
  • Gloves
  • Lab coat
  • Goggles
  • Vacuum equipment
  • Glass scintillation vials
  • Glass Pasteur pipettes
  • Embedding mould
  • 60C oven
  • Gelatin capsules


Transmission electron microscopy (TEM)



Fresh specimens are collected and cut/dissected into 2mm2pieces as soon as possible after collection. The specimens should be cut/dissected whilst under the aldehyde fixative, then placed in small glass vials filled with fresh fixative. Lids should be kept on the vials. Vials should be kept slowly rotating (<4 rpm) during the entire procedure. If there is a problem with the samples floating on the surface of the aldehyde fixative then keeping the samples in the fixative in open topped vials under vacuum may help. Keep the samples under vacuum for 15 minutes, then release and shake the vials. If samples are still floating repeat the process. The procedures below are general protocols and may need to be adapted for each sample. Aldehyde fixation times of between 1 and 24 hours are not unknown and reference should be made to the literature for similar material. Temperature (e.g., 4 or 20C) and osmolarity may play a role in good fixation. The addition of 0.1M sucrose (osmolarity = 1093mOsmols ) may help with particular samples. With all the protocols listed below the amount of time the specimens are in resin depends on the particular sample. As with the fixation reference should be made to the literature for similar material. Five changes of resin over a 24 to 72 hour period, to ensure complete penetration are not uncommon. Another technique is to keep the samples in the resin, for three hours, with fresh resin changes every hour, on a rotator, at 40C in the oven, so that the resin becomes less viscous and can penetrate better. Evacuation of the samples can also aid in resin penetration.

Procedure 1

No. Treatment Time Temp.
1. Aldehyde fixative 3 hours 25C
2. Buffer Wash 5 min 25C
3. Buffer Wash 5 min 25C
4. Osmium 2 hours 5C
5. Buffer Wash 5 min 5C
6. Buffer Wash 10 min 5C
7. Buffer Wash 10 min 5C
8. Buffer Wash 30 min 5C
9. 30% EtOH 5 min 5C
10. 50% EtOH 10 min 5C
11. 70% EtOH 30 min 5C
(Leave overnight in hand fixation schedule)
12. 90% EtOH 30 min 5C
13. 100% EtOH 30 min 5C
14. 100% EtOH 30 min 5C
15. 100% EtOH 1 hr 5C
16. 1:3 resin:EtOH 1 hr 5C
17. 1:1 resin:EtOH 1 hr 5C
18. 3:1 resin:EtOH 1 hr 5C
19.(to be repeated 3 times) 100% resin 1 day 5C

Specimens are transferred individually into gelatin capsules capsules, supported by the edges of holes cut into the lids of TEM film boxes, for polymerisation. The lids of the capsules need not necessarily be used but if they are then holes must be pierced into them to allow pressure equilibration under vacuum. The resin is polymerised, between 58-60C at 440mmHg, for 18 to 24 hours. Blocks should be left for 2 to 3 hours, in the fume cupboard, before being sectioned, to allow the resin to harden. Gelatin capsules can be dissolved away in warm water, however, this is not essential. Blocks can be individually coded using an indelible marker pen and logged for future reference. Alternatively, specimens can be flat embedded, cut out of the polymerised disk of resin, reoriented and glued to a resin stub for sectioning.

Procedure 2

No. Treatment Time Temperature
1. Aldehyde Fixative 3 hr 5C
2. Buffer 30 min 5C
3. Osmium 2 hr 5C
4. Buffer 10 min 8C
5. Buffer 10 min 11C
6. Buffer 10 min 14C
7. 30% EtOH 30 min 17C
8. 50% EtOH 30 min 20C
9. 70% EtOH 45 min 20C
10. 90% EtOH 1 hr 20C
11. 100% EtOH 1 hr 20C
12. 100% EtOH 1 hr 20C
13. 1:3 resin:EtOH 1 hr 17C
14. 1:1 resin:EtOH 1 hr 30 min 14C
15. 3:1 resin:EtOH 2 hr 11C
16. 100% resin 2 hr 8C
17. 100% resin 4 hr 5C
18. 100% resin 4 hr 5C
19. 100% resin 4 hr 5C
20. 100% resin 4 hr 5C

Polymerisation of the specimens as above.


There are various types of resins available for TEM embedding. Resins may vary considerably with respect to viscosity and grades of hardness, which are the main criteria for selecting a resin. Generally, it is recommended to try out several embedding approaches and resins when working with a new tissue. The user can then find out which resin works best for that specific tissue, although this can also be a matter of personal taste.

LR White (London Resin Company) has become a popular and generally used resin because of its low viscosity and fairly rapid infiltration of most types of botanical tissue. LR White resin polymerizes at low temperature using UV irradiation at 365 nm, but also at higher temperatures (ca. 60C) when using an oven. Recommended polymerization time is overnight. LR Gold is an alternative resin that is most useful for embedding unfixed material at low temperatures (i.e. < 1C). A major advantage of LR White resin is the hydrophilic nature of the sections, which makes this resin widely used for immunolabelling. Compared to Epon resin sections, however, LR White sections may show more wrinkles and bumps. Since the toxicity of LR White has not been studied in detail, it is recommended to treat this resin as toxic.

Another frequently used resin is Spurr resin, which is a typically hard grade resin. This resin has an excellent penetration capacity for tough materials such as lignified plant tissue and hard specimens. However, Spurr resin should be treated as a suspect carcinogen, which is the main reason why some laboratories forbid its use. Curing of Spurr resin can be done at 70C for ca. 8 hours.

Alternative resins include Epon, Araldite, Unicryl, and TAAB resin. However, since the viscosity for most of these resins is higher than for Spurr and LR White, they may cause infiltration problems with some tissues. It is also possible to experiment with mixtures of resins, such as Mollenhauer’s Epon/Araldite resin.

Various tips for working with resins (based on Glauert 1991)

  1. Store the basic resin components at room temperature rather than in the refrigerator.
  2. Make up a fresh resin mixture for each embedding session rather than re-using an old mixture. Storing complete mixture in the fridge may increase viscosity and result in infiltration problems.
  3. It is usually a good idea not to exceed 60C for polymerization of resins as this may result in more consistent blocks.
  4. Moisture may adversely affect resin accelerators. Therefore, keep resin bottles in dry conditions, write the date on the bottle when first used, and discard the resin after it has been open for more than 6 months. Keep also an eye on the expenditure date.
  5. Use a reliable text book for the preparation of resins since various errors exist in literature.
  6. Health and safety: use gloves and work under the fume cupboard. Avoid contact with the complete resin and its components and minimize exposure to dust or chips during block trimming or sectioning. It is recommended to use a vacuum cleaner to remove the trimming debris continuously while you work. Always wash your hands immediately after working with resins.



Embedded samples can be trimmed with an automatic specimen trimmer. Alternatively, resin blocks are fixed in a holder underneath a dissecting scope and trimmed manually with a sharp single edge industrial razor blade. First, expose the specimen by making a horizontal cut across the top of the block. Then, trim the sides of the block creating a -pyramid’ with a trapezoid shape face at the top. The resins can have adverse effect on human health; therefore, avoid breathing the resin dust and keep your workspace clean as much as you can.


Sectioning is done with an ultramicrotome (e.g. Reichrt-Jung Ultracut E, Leica UltraCut E Microtome).

Sectioning procedure summary:

  1. Turn on the microtome illumination. Tightly clamp the resin block into a specimen holder and attach the knife to the knife holder. Typically, the knife angle should be around 5°.
  2. Bring the block and knife close together so that you can see both the block face and the knife edge with a binocular magnifier of ultramicrotome
  3. Focus the magnifier on the knife edge.
  4. Align the block face with the knife edge (for more information how to do that see the detailed description below).
  5. Make sure that you use the correct settings for the cutting stroke and suitable cutting speed.
  6. Advance the block face towards the knife and observe the reflection of the knife edge getting thinner and thinner. Use the 2μm and 1μm advance once you get very close to the knife edge.
  7. Once you are not able to see the reflection switch to 0.2μm advance and watch for the first section.
  8. As soon as the block face touches the knife and you get the first (most likely incomplete) section, stop the microtome advance and fill up the knife boat with fresh distilled water. Swipe the surface of the water with a clean lens paper to remove any dust particles and debris. Adjust the water level so that the water reaches the knife edge (but not more than that).
  9. Switch to the ultrathin sectioning mode and cut sections ideally between 60 and 90nm thick. The sections will float on the surface of water and their thickness can be determined based on their color. Once sections are being cut it may be necessary to add a few small drops of water so that the section color can be observed and so that the sections float freely away from the knife edge. Silver and gold sections are ideal for the transmission electron microcopy.

More detailed description of the sectioning procedure, tips for mastering the specimen-knife alignment:

Place the resin block into a specimen holder and mount the knife in the knife holder. Adjust the settings for the cutting stroke. The specimen arm movement has an elliptical trajectory and variable speed during the individual cutting cycle. The arm should be closer to the knife when moving down and return back up at the beginning of the next cutting stroke further from the knife. The specimen arm should move at the slowest speed when the block face moves past the knife edge (i.e. when the sections are being cut). It is also important to adjust the amplitude of the arm movement base on the height of the knife you use. Carefully bring the specimen closer to the knife so that you can observe both the block face and the knife edge with the binocular magnifier. Focus on the knife edge. Now you are ready to start with a finer adjustment of the knife and specimen. Proper alignment of knife and specimen block is critical for successful sectioning and requires practice and patience. You can change angles and positions of your block and knife using the knobs on the microtome (see the manual of your specific microtome). If new to ultrathin sectioning, practicing with a blank resin block before sectioning the real sample may be helpful. First, the parallel sides of the trapezoid block face are oriented parallel to the knife edge (see Fig.1 front view). Subsequently, the surface of the block face must be oriented parallel to the knife edge in both vertical and horizontal directions (Fig.1 top and side view).

Fig.1: Aligning the resin block and knife for thin sectioning. Possible correct and incorrect positions of the block with respect to the knife edge in all three dimensions.

The proper alignment is judged by observing the reflection of the knife edge on the block face. The width of the reflection represents the distance between the knife and the block face. If the knife and the block face are parallel to each other, the reflection should have the same thickness across the block face (Fig. 1 top view) and the same thickness should be maintained when the block moves past the knife edge (Fig. 1 side view). When reflection disappears the knife and block face are touching (or almost touching).

Although comprehensive descriptions of how to achieve correct alignment exist in the literature, practicing with your specific material and equipment is priceless. The reflection may appear somewhat different depending on the specific microtome, knife and resin you use. Blank resin blocks (i.e. without the sample) are great for learning as the face of a blank block provides very clear and shiny reflection. In order to figure out how the reflections indicating correct alignment between knife and specimen should look like, observing the change in reflection while the specimen is slowly retreated might be helpful. First, perform a coarse alignment of the specimen (use a non-important trial sample or a blank resin block) and trim back the block face with a glass knife. Now your block face should be parallel to the knife edge. Cut several semi -thin (0.1-0.5μm) sections and if satisfactory, retreat the block from the knife edge by turning the advance knob backwards and observe the change on the block face. If the illumination system of your microtome is working properly, you should see the reflection getting thicker and thicker as the block is moving further apart from the knife and the thickness of the reflection should not change when block face is moved up and down past the knife edge. Try reorienting the sample from its parallel position and observe how the reflection changes (the reflection is not uniformly thick). Then you can attempt to align the block back to be parallel to the knife, advance it towards the knife edge and cut a few sections. Once you feel comfortable doing the alignment you can cut ultrathin sections using a fresh glass knife or a diamond knife. Use a diamond knife with great care and only if you are thoroughly familiar with the sectioning procedure. Diamond knives are very sensitive and very expensive.

Some most common factors causing problems with ultrathin sectioning:

  1. Lack of practice, incorrect alignment between block face and knife edge ®practice more, use blank resin block to practice
  2. The block face is too big ®trim the block face smaller (creating a triangular instead of trapezoid block face can make sectioning of a very tough material easier)
  3. The material you are trying to cut has internal structure and you are cutting it “against the grain” (e.g. when making cross-sections of wood tissue, cut along the rays, not perpendicularly) ®turn/reorient the sample and try to section it again
  4. Too low/ too high level of water in the knife boat (the sections are not floating smoothly off the knife edge/the block face is picking up water) ®adjust the water level accordingly
  5. Your knife is dull (compressed sections, irregular sectioning) ®change the knife or move to a different part of the knife edge
  6. Poor infiltration of the sample, incomplete polymerization of the resin ®prepare new samples

Picking up sections

Collecting sections on grids is another -tricky’ step that requires practice. If you are lucky to have ribbons of sections rather than individual sections, picking them up should be easier. The easiest way how to pick sections up is by touching the floating sections from the top with a grid. This method is especially convenient for coated grids that are difficult to immerse under water. A small drop of water holding the sections will attach to the grid surface and once the water evaporates away, the sections will be mounted firmly onto the grid. However, this method of picking up sections has several drawbacks. First, it takes about 10-15min before the drop evaporates. Keep the grid undisturbed still attached to the forceps during that time (use clamps to hold the forceps arms together). Second, the sections sometimes tend to slide down from the convex drop and attach to the margin of the grid. Therefore, keep the grid leveled when waiting for the drop to evaporate. In addition, wrinkles may sometimes appear on the sections. Alternative method for picking up sections is to immerse the grid vertically under the water level, move it close to the sections and then pull it up. Nice and smooth sections should adhere to the grid. However, this method requires a still hand and a lot of practice.


Staining can be conducted with uranyl acetate and lead citrate using an automatic staining apparatus or can be done manually. These stains are by far the most widely used general counterstains that react both with negatively and positively charged side-chains of proteins.

4% uranyl acetate in distilled water

Weigh out 2 gm of uranyl acetate.

Transfer to amber glass bottle.

Add 50 ml of distilled water.

Add a small stir bar and stir for at least 1 to 2 hours.

Let settle overnight. Store in darkness.

Filter a small amount immediately before use.

Reynolds’ lead citrate

  • Prepare fresh 1N NaOH in advance: 2 gm of NaOH pellets into 50 ml of distilled water
  • Allow time for this to dissolve and mix.
  • Weigh out 1.33gm of lead nitrate.
  • Weigh out 1.76 gm of sodium citrate.
  • Add the above to about 30 ml of distilled water in volumetric flask or cylinder with stopper. Stopper and shake vigorously for one minute and intermittently for 30 minutes. The solution should be milky in appearance.
  • Add 8 ml of the 1N NaOH solution to the above solution; mix gently and then bring volume to 50 ml with distilled water. Stopper and mix by inversion. The staining solution should now be clear.
  • Lead Citrate stain is stable for approx. 6 months.

Manual staining:

Staining is performed manually by floating grids on drops of stains. Fresh drops of stain are prepared on a Parafilm placed in a Petri dish just before using them. First, float the grid on a drop of uranyl acetate for 15-30min (depends on your sample, resin you used). The staining should be conducted in dark (e.g. in a Petri dish covered with an aluminum foil). Then, grids are washed thoroughly with distilled water. To do that hold the grid vertically above a beaker with forceps and gently jet wash the grid with water using a squeeze bottle (let the stream/drops of water run in between the arms of the forceps down over the grid). Drain the excess water onto a filter paper and suck off the water trapped in between the forceps arms otherwise the grid will stick to the forceps due to the surface tension. Then, place the grid on a drop of lead citrate for 5-10min. Lead citrate precipitates with CO2. To absorb CO2 from the air in the dish, it is recommended to put few NaOH pellets around the Parafilm with the stain and wet them with a few drops of water. After staining, wash the grids with distilled water as described previously. Keep the grids away from dust in a small Petri dish or a special grid storage box until observed.


  1. Specimens are fixed in 3% glutaraldehyde in 0.5 M cacodylate buffer at pH 6.8 to 7.2 for 1.5 hrs and washed 3 hrs in buffer.
  2. Samples are incubated in 40% methylamine for between 24 and 108 hours before dehydration and embedding.


Toluidine blue is a general stain that binds to virtually all areas of the cell. Thus walls, cytoplasm, vacuoles and nuclei are all rendered visible. This stain is metachromatic and stains different cellular components different colors. Toluidine blue binds to acidic polyanions.

  1. Prepare 0.5% toluidine blue O in 0.1M phosphate buffer, pH 7.0. Filtering the solution is recommended.
  2. Cut semi-thin sections and mount on glass slides.
  3. Dip slides in the stain for 1-10 min depending on type of specimen.
  4. Wash slide in distilled water.


  1. Dissolve 0.25g of powdered Formvar in 50ml chloroform (cover beaker with parafilm) with vigorous stirring, to give 0.5% Formvar solution. Make fresh every time.
  2. Rinse glass slides with 100% ethanol and air dry.
  3. Dip slides into mixture and stand upright on tissue to drain. While draining and drying, keep the slide isolated from dust and air currents.
  4. Fill a large dish/trough brimful with distilled water and clean the surface by wiping with Velin tissue.
  5. Use the tips of a pair of forceps to score around the edges of the film on a dipped slide. Ensure that the scoring is continuous.
  6. Breathe on the slide to facilitate film separation.
  7. Float off the Formvar film by gently easing the slide under the water surface at a moderate angle: this can be monitored by viewing the surface at an appropriate angle.
  8. Either immerse the slide completely and release; or withdraw it gently after the film is seen to float free.
  9. Drop grids onto the floating film, shiny side up.
  10. Retrieve with a sheet of filter paper and allow to dry.



  1. Phosphate Buffer (PB):a. Mix 40.5ml 0.2M Na2HPO4stock solution (X solution) with 9.5ml 0.2M NaH2PO4stock solution (Y solution).b. Dilute to 100ml in a volumetric flask with distilled H2O to 0.1M final conc., pH 7.4.
  2. Phosphate Buffer Saline Tween (PBST):To 20ml of PB add 80ml distilled H2O, 0.9g NaCl, 100μl Tween 20 and 0.02g Na-azide.This gives a 20mM solution of phosphate buffer with 0.9% NaCl, 0.1% Tween 20 and 0.02% Na-azide.

Other Solutions

  1. 10% dried milk in distilled H2O centrifuged for 10 – 20 mins. Use the supernatant to make up solution 2 below.
  2. 3% dried milk – diluted from solution 1.
  3. 5% (v/v) H2O2in distilled H2O.


Temp: 25 – 27C Controls: Mosaic virus, D-2, -glucosidase or blank

  1. Ultrathin sections are collected onto nickel (do not use copper) grids.
  2. Reaction wells are created by depressing the broader end of a 1ml Eppendorf pipette tip on a piece of parafilm.In order to minimize evaporation during each reaction, a moisture chamber should be made by covering each well with a plastic lid.
  3. Sections of osmicated material are etched with 5% H2O2 for 5 – 10 mins.
  4. Wash twice in PBST. (5 – 10 mins)
  5. Block in 10% aqueous low fat dried milk for 1 hr.
  6. Incubate for 1 hr in, for example, rabbit pectinase antibody 1 : 1500 diluted in PBST containing 0.1% Tween 20 and 3% low fat dried milk.
  7. Wash 3x in PBST. (5 – 10 mins)
  8. Block in 10% low fat dried milk for 30 mins.
  9. Incubate for 1hr in, for example, goat anti-rabbit IgG conjugated to 10 nm gold particles (Sigma), diluted 1 : 50 in PBST containing 3% low fat dried milk.
  10. Wash twice in PBST for 10 mins each.
  11. Wash once in d.H2O for 10 mins.
  12. Fix in 2% glutaraldehyde in d.H2O.
  13. Wash twice in distilled H2O and dry.
  14. Sections are contrasted with uranyl acetate and lead citrate.


After washing in 0.05 M phosphate buffer, samples are stained with ruthenium red to localize acidic pectins.

Samples stained with ruthenium red are fixed in Karnovsky’s fixative containing 0.1% ruthenium red for 1.5 h and washed again in buffer containing 0.1% ruthenium red. These samples are then postfixed in 1% buffered OsO4 containing 0.1% ruthenium red for 1.5 h at room temperature. A brief buffer rinse is followed by dehydration and embedment in resin.


Samples for the hydroxylamine-ferric chloride reaction are prepared by postfixation in 1% buffered OsO4. After washing in phosphate buffer, H-samples are dehydrated in a 25% and then 60% ethanol-water solution for 15 min each. Subsequently, they are put for 20 min in a fresh 60% ethanol-water solution containing 14% hydroxylamine hydrochloride and 14% NaOH (w/v). H-samples are then washed for 5 min in 0.1 M HCl in 60% ethanol, stained for 20 min in a fresh solution of FeCl3 (20% w/v in 60% ethanol), and washed in 60% ethanol.


PATAg (periodic acid-thiocarbohydrazide-silver proteinate) staining should be conducted using non-osmicated material because of the possibility that osmium can interfere with the PATAg reaction. Ultra-thin sections from non-osmicated samples are collected on 100 mesh gold grids and floated for 30 min on 1% aqueous periodic acid, washed three times with distilled water, and floated for 2 h on 0.2% thiocarbohydrazide in 20% acetic acid. The sections are then washed for 10 min in 10% acetic acid, followed by 2 washes in 4% and 2% acetic acid for 5 min in total, and three washes of 5 min in distilled water. Finally, the grids are floated for 30 min on 1% aqueous silver proteinate solution in absolute darkness. For control, either the periodic acid-oxidation or the thiocarbohydrazide-incubation step should be omitted.


  1. Prepare a 0.2M solution of dibasic sodium phosphate withNa2HPO4 – 28.39gor Na2HPO4.2H2O – 35.61g

    or Na2HPO4.7H2O – 53.65g

    or Na2HPO4.12H2O – 71.64g

    and distilled water to make 1000ml.

  2. Prepare a 0.2M solution of monobasic sodium phosphate withNa2H2PO4.H2O – 27.6gor Na2H2PO4.2H2O – 31.21g

    and distilled water to make 1000ml.

  3. Prepare the 0.1M phosphate buffer by mixing x ml of 0.2 ml of 0.2M dibasic sodium phosphate with y ml of 0.2M monobasic sodium phosphate and diluting to 100 ml with distilled water.
pH (at 25C) x ml y ml
5.8 4.0 46.0
6.0 6.15 43.85
6.2 9.25 40.75
6.4 13.25 36.75
6.6 18.75 31.25
6.8 24.5 25.5
7.0 30.5 19.5
7.2 36.0 14.0
7.4 40.5 9.5
7.6 43.5 6.5
7.8 45.75 4.25
8.0 47.35 2.65

The osmolarity of the buffer is adjusted by varying the molarity of the phosphates, or by the addition of sucrose, glucose or sodium chloride. The osmolarity of 0.1M Sörensen’s buffer at pH 7.2 is 226mOsmols (Fahimi and Drochmans 1965b); corresponding values for 0.05M, 0.075M and 0.15M buffers are 118, 180 and 350mOsmols , while addition of 0.18M sucrose to a 0.1M buffer raises the osmolarity to 425mOsmols .



  • 50ml 0.1M phosphate buffer, pH 7.2
  • 12ml 25% glutaraldehyde in water
  • 38ml distilled water.
  • Note: if glutaraldehyde is added to the buffer before water is added, a precipitate might form.


  1. Prepare 0.1M phosphate buffer, pH 7.2.
  2. Prepare 20ml of a 10% solution of paraformaldehyde by dissolving 2.0g of paraformaldehyde powder in 20ml of distilled water and heating to 60-65C. Add a few drops of 1.0M sodium hydroxide until the solution clears. Allow the solution to cool before use.
  3. Prepare the fixative with:
    • 0.1M phosphate buffer – 50ml
    • 10% paraformaldehyde solution – 20ml
    • 25% glutaraldehyde solution – 10ml
    • distilled water – 20ml

This fixative contains 2% paraformaldehyde and 2.5% glutaraldehyde in 0.05M phosphate buffer.
Note: 0.05% Ruthenium red can be added in order to contrast the middle lamella of the cell wall.


  1. Osmium tetroxide is purchased in 5ml quantities of a 2% solution in closed vials and kept in the fridge.
  2. Snap off the top carefully in the fume cupboard. Using a Pasteur pipette, remove the osmium tetroxide solution from the vial into a glass stoppered mixing flask. The flask should ideally be brown as the solution is light sensitive. However, if none is available then a clear glass bottle may be used if it is surrounded by aluminium foil.
  3. Add 5ml of 0.1M phosphate buffer. Shake gently to mix. The pipette and empty vial should be disposed of into a waste osmium tetroxide bottle.

Osmium tetroxide is also purchased in 2ml quantities of a 4% solution. The procedure is as above but you mix 2ml OsO4 with 2ml distilled water and 4ml of 0.1M phosphate buffer.

Also available are 0.1g crystals of osmium which need to be dissolved in 10 ml of 0.05M phosphate buffer. Again the precautions are as above.


See 13.2. (above)


For related protocols, see:
Tissue Preparation for Transmission Electron Microscopy
Preparation of Material for SEM Examination
Cryo-SEM and quantitative cryo-analytical SEM


Chaffey N.J. 2002. Conventional (chemical-fixation) transmission electron microscopy and cytochemistry of angiosperm trees. In N. J. Chaffey (Ed.), Wood formation in trees: cell and molecular biology techniques, 41-64. Taylor & Francis, London, UK.

Glauert A.M. 1991. Epoxy resins: an update on their selection and use. Microsc. and Analysis (September): 15-20.

Gluaert A.M., Lewis P.R. 1998. Biological specimen preparation for transmission electron microscopy. Volume 17: practical methods in electron microscopy. A.M. Glauert (Ed.), Portland Press, London and Miami.


Chloroform – Very toxic by inhalation, causing drowsiness, nausea, vomiting and unconsciousness. Toxic by ingestion. irritating to skin and eyes, possibly causing conjunctivitis and burning. Has been found to cause cancer in lab animals. May cause adverse mutagenic or teratogenic effects.

Ethanol – Intoxicating if inhaled or ingested. Irritating to eyes. If ingested in undiluted form has a severe drying effect on mucous membranes of mouth and throat. Can be damaging if splashed in eyes. Highly flammable. Vapour/air mixture explosive. Can react vigorously with oxidising materials. Can react violently with potassium oxides and potassium. Ignites in contact with platinum-black catalyst. Can react violently with silver nitrate.

Formvar – Oxidising agent. Harmful in contact with skin, eyes and if swallowed. Avoid inhalation of dust.

Glacial acetic acid – Vesicant. Causes severe burns. Causes internal irritation and damage if taken by mouth. Irritating vapour. Flammable. Vapour/air mixture explosive. Can react vigorously with oxidising materials. Dangerous in contact with chromic acid, sodium peroxide and nitric acid. Causes vigorous exothermic polymerisation of acetaldehyde. Vapour can ignite in contact with potassium.

Glutaraldehyde – Harmful by ingestion and inhalation. Extremely irritating to eyes. Prolonged skin contact can cause dermatitis and sensitisation. Evidence of reproductive effects. Combustible. Can react vigorously with oxidising materials.

Hydrogen peroxide – Extremely irritating to eyes and respiratory system. Irritating to skin. May cause burns if contact is prolonged. If ingested, sudden evolution of oxygen may cause injury by acute distension of the stomach, and may cause nausea, vomiting and internal bleeding.

Lead citrate – Toxic by inhalation and if swallowed. Danger of cumulative effects.

LR white resin – harmful by inhalation, in contact with skin and if swallowed. Risk of serious damage to eyes. Irritating to respiratory system and skin. May cause sensitisation by skin contact. Emits toxic fumes under fire conditions.

Methylamine – Irritating to eyes, respiratory system and skin.

Na-azide – Very toxic if swallowed. Contact with water liberates toxic gas. Contact with acid liberates very toxic gas.

Osmium tetroxide – May be fatal if inhaled, swallowed or absorbed through the skin. Causes severe irritation. High concentrations are extremely destructive to tissues of the mucous membranes and upper respiratory tract, eyes and skin. Symptoms of exposure may include burning sensation, coughing wheezing, laryngitis, shortness of breath, headache, nausea and vomiting. Lab experiments have shown mutagenic effects. Contact with combustible material may cause fire.

Paraformaldehyde – Harmful by ingestion. Irritating to skin, eyes and respiratory system. Evidence of mutagenic effects. Flammable. May evolve toxic fumes in a fire. Can react vigorously with oxidising materials. Any formaldehyde evolved can react with hydrochloric acid to produce a carcinogenic compound.

Periodic acid – Causes severe burns to eyes and skin. If ingested causes severe internal irritation and damage. Vapour is irritating and corrosive to eyes and respiratory system.

Phosphate buffer – Sodium dihydrogen orthophosphate and di-sodium orthophosphate may irritate eyes and respiratory system if inhaled as dust. Ingestion of large amounts of phosphate can cause serious disturbances in calcium metabolism. May evolve toxic fumes in a fire.

Resins – Use gloves and work under the fume cupboard. Avoid contact with the complete resin and its components and minimize exposure to dust or chips during block trimming or sectioning. It is recommended to use a vacuum cleaner to remove the trimming debris continuously while you work. Always wash your hands immediately after working with resins.

Sodium cacodylate – Toxic by inhalation and if swallowed. Danger of cumulative effects. Gaseous decomposition products are extremely toxic.

Sodium chloride – Ingestion of large amounts may cause diarrhoea, nausea, vomiting, hypernoea and convulsions. May irritate eyes.

Sodium dihydrogen orthophosphate and di-sodium hydrogen orthophosphate – May irritate eyes and respiratory system if inhaled as dust. Ingestion of large amounts of phosphate can cause serious distubances in calcium metabolism.

Sucrose and glucose – Ingestion of significant quantities may lead to metabolic imbalances. May irritate eyes. Combustible. May react explosively in intimate mixtures with oxidising materials. May be explosive if finely divided material is dispersed in air.

Thiocarbohydrazide – very toxic by inhalation, in contact with skin and if swallowed. Heating may cause an explosion.

Toluidine blue – Harmful.

Tween – Possibly harmful.

Uranyl acetate – Toxic by inhalation and if swallowed. Danger of cumulative effects.


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